In the relatively short time since gene editing involving clustered regularly interspaced short palindromic repeats (CRISPR) arrived on the life-sciences scene—most particularly in the handful of years since we got CRISPR/Cas9 and a much more simplified editing process—the technology has seen its ups and downs with regard to how safe, specific, efficient and reliable it is. But there is no doubt the technology continues to advance and almost certainly will hold a key position in the genomics arena for a long time to come.

Germany’s Merck KGaA (not to be confused with U.S.-based Merck & Co.), for example, recently developed an alternative CRISPR genome editing method that it says makes CRISPR “more efficient, flexible and specific, giving researchers more experimental options and faster results that can accelerate drug development and access to new therapies.”

Merck KGaA calls the new technique proxy-CRISPR and maintains that it provides access to previously unreachable areas of the genome. Most natural CRISPR systems, found in bacteria, cannot work in human cells without significant re-engineering, the company notes; however, proxy-CRISPR is said to provide a simpler and quicker method to increase their usability without the need to re-engineer native CRISPR proteins.

“With more flexible and easy-to-use genome-editing technologies, there is greater potential in research, bioprocessing and novel treatment modalities,” said Udit Batra, a member of the company’s executive board and CEO of its Life Science unit. “As a leader in genome editing, Merck’s new technology is just one example of our commitment to solving challenges in the genome editing field, and we will continue to make CRISPR research a priority.”

The new technology is a follow-on to Merck’s existing CRISPR applications, and the company’s next suite of genome-editing tools for the research community—planned for launch later this year—is expected to include novel and modified versions of Cas and Cas-like proteins.

More progress on the RNA front

Researchers in the medical school at the University of California, San Diego (UC San Diego) in a 2016 study repurposed the CRISPR/Cas9 technique to track RNA in live cells in a method called RNA-targeting Cas9 (RCas9). In a new study, published Aug. 10 in Cell, the team took RCas9 a step further, using the technique to correct molecular mistakes that lead to microsatellite repeat expansion diseases, which include myotonic dystrophy types 1 and 2, the most common form of hereditary amyotrophic lateral sclerosis and Huntington's disease.

“This is exciting because we’re not only targeting the root cause of diseases for which there are no current therapies to delay progression, but we’ve re-engineered the CRISPR/Cas9 system in a way that’s feasible to deliver it to specific tissues via a viral vector,” said senior author Dr. Gene Yeo, professor of cellular and molecular medicine at UC San Diego School of Medicine.

Microsatellite repeat expansion diseases arise because there are errant repeats in RNA sequences that are toxic to the cell, in part because they prevent production of crucial proteins. These repetitive RNAs accumulate in the nucleus or cytoplasm of cells, forming dense knots, called foci.

In this proof-of-concept study, Yeo’s team used RCas9 to eliminate the problem-causing RNAs associated with microsatellite repeat expansion diseases in patient-derived cells and cellular models of the diseases in the laboratory.

There is still a ways to go before RCas9 could be tested in patients, though, Yeo acknowledged. One bottleneck is efficient delivery of RCas9 to patient cells, as the non-infectious adeno-associated viruses that are commonly used in gene therapy are typically too small to hold Cas9 to target DNA. Yeo’s team made a smaller version of Cas9 by deleting regions of the protein that were necessary for DNA cleavage, but dispensable for binding RNA.

“The main thing we don’t know yet is whether or not the viral vectors that deliver RCas9 to cells would illicit an immune response,” he said. “Before this could be tested in humans, we would need to test it in animal models, determine potential toxicities and evaluate long-term exposure.”

Reducing off-target effects

Integrated DNA Technologies (IDT) in early August announced the launch of what it says is the first Cas9 enzyme variant that extensively reduces off-target effects in CRISPR genome editing without compromising on-target activity. The Alt-R S.p. HiFi Cas9 Nuclease 3NLS enzyme is a recombinant S. pyogenes Cas9 mutant that improves specificity while maintaining a high editing efficiency similar to wild-type Cas9, the company explains, adding that the launch represents a “major step towards therapeutic use of CRISPR, which has previously borne the risk of the unwanted off-target editing events observed with wild-type Cas9.”

Earlier Cas9 mutants that offer improved specificity suffer from a moderate to severe loss of on-target activity when used as a ribonucleoprotein (RNP) complex. As IDT explains, “the largely preferred method of delivering genome-editing reagents as RNP complexes reduces, but does not eliminate, the risk of off-target editing. However, recent attempts at rational design of Cas9 mutants with reduced off-target activity traded on-target activity for improved specificity, and produced mutants generally unsuitable for use in RNP delivery.”

In order to successfully provide a Cas9 mutant with radically reduced off-target effects while maintaining high on-target activity, IDT screened more than 250,000 mutants in two rounds of selection.

“We performed an unbiased evaluation of several versions of high fidelity Cas9 enzyme in primary human stem cells. We have been very impressed with the characteristics of this new IDT enzyme,” said Dr. Matt Porteus, of Stanford University’s Division of Stem Cell Transplantation and Regenerative Medicine in a recent IDT news release. “Unlike other versions, this version consistently gives us high on-target editing activity while having low off-target activity. Because of the retained excellent on-target activity and improved specificity profile, we are excited to use this version in our future experiments focused on developing novel genome editing-based therapies for severe diseases with unmet medical needs.”

Improving CRISPR/Cpf1

Scientists on the Florida campus of The Scripps Research Institute (TSRI) have improved and broadened the ways a “competitor” to CRISPR/Cas9—the CRISPR/Cpf1 editing system—may be used to study and fight human diseases. To achieve this, TSRI professor Michael Farzan, co-chair of TSRI’s Department of Immunology and Microbiology, and TSRI research associate Guocai Zhong incorporated into the CRISPR-Cpf1 guide RNAs with “multiplexing” capability, meaning multiple genetic targets in a cell may be hit by each CRISPR/Cpf1 complex.

“This system simplifies and significantly improves the efficiency of simultaneous editing of multiple genes, or multiple sites of a single gene,” Zhong said. “This could be very useful when multiple disease-related genes or multiple sites of a disease-related gene need to be targeted.”

“This approach improves gene editing for a number of applications,” Farzan added. “The system makes some applications more efficient and other applications possible.”

The study was published as an advanced online paper in the journal Nature Chemical Biology on June 19, 2017.

Over the last five years, TSRI notes, the CRISPR gene-editing system has “revolutionized microbiology and renewed hopes that genetic engineering might eventually become a useful treatment for disease. But time has revealed the technology’s limitations. For one, gene therapy currently requires using a viral shell to serve as the delivery package for the therapeutic genetic material. The CRISPR molecule is simply too large to fit with multiple guide RNAs into the most popular and useful viral packaging system.”

The new study from Farzan and colleagues helps solve this problem by letting scientists package multiple guide RNAs.

This advance could be important if gene therapy is to treat diseases such as hepatitis B, Farzan said. After infection, hepatitis B DNA sits in liver cells, slowly directing the production of new viruses, ultimately leading to liver damage, cirrhosis and even cancer. The improved CRISPR-Cpf1 system, with its ability to multiplex, could more efficiently digest the viral DNA before the liver is irrevocably damaged, he said.

“Efficiency is important. If you modify 25 cells in the liver, it is meaningless. But if you modify half the cells in the liver, that is powerful,” Farzan said. “There are other good cases—say, muscular dystrophy—where if you can repair the gene in enough muscle cells, you can restore the muscle function.”

Two types of these molecular scissors are now being widely used for gene editing purposes: Cas9 and Cpf1. Farzan said he focused on Cpf1 because it is more precise in mammalian cells. The Cpf1 molecule they studied was sourced from two types of bacteria, Lachnospiraceae bacterium and Acidaminococus sp., whose activity has been previously studied in E. coli. A key property of these molecules is they are able to grab their guide RNAs out of a long string of such RNA, but it was not clear that it would work with RNA produced from mammalian cells. Guocai tested this idea by editing a firefly bioluminescence gene into the cell’s chromosome. The modified CRISPR-Cpf1 system worked as anticipated.

“This means we can use simpler delivery systems for directing the CRISPR effector protein plus guide RNAs,” Farzan said. “It’s going to make the CRISPR process more efficient for a variety of applications.”

Looking forward, Farzan said the Cpf1 protein needs to be more broadly understood so that its utility in delivering gene therapy vectors can be further expanded.

Commentary: Two barriers to CRISPR’s success and how they can be overcome

One of CRISPR’s most attractive benefits is its ability to scan the genome for a particular genetic sequence and make a customized edit, as if the genome was a line of text on a computer screen. When it first arrived on the molecular biology scene, researchers celebrated CRISPR’s flexibility and potential for treating genetic illness. Recently, however, molecular biologists have expressed concern over two of CRISPR’s aspects: (1) its low specificity, or its tendency to introduce mutations where it is not supposed to, and (2) its low efficiency, its relative inability to perform edits when it is supposed to.

Here, we will discuss these two concerns as well as the role of two novel molecular techniques, DNA-free CRISPR and heteroduplex assays, in reducing the impact of CRISPR’s limitations.

High prevalence of unintended mutations

A recent commentary on CRISPR (“CRISPR—The past, present and future” by Dr. Adriano Flora of Taconic Biosciences in the May 2017 issue of DDNews) suggested that researchers needn’t concern themselves over CRISPR’s potential for producing non-specific effects. But the author left open the possibility that new data could change this perception. Indeed, shortly after that commentary was published, a letter published online in Nature Methods on May 30 titled “Unexpected mutations after CRISPR-Cas9 editing in vivo” described the vast extent to which CRISPR produced unintended mutations in the mouse genome. As it turns out, CRISPR’s tendency to introduce unintended mutations into the genome of a cell is one of its most significant flaws.

After using CRISPR to treat blindness in mice, a group of researchers sequenced the animals’ entire genomes, searching for all possible mutations down to the single-nucleotide level. Subsequently, the researchers discovered more than 1,500 single-nucleotide mutations and more than 100 large-scale insertions and deletions in their mice. This study introduced the possibility that CRISPR could potentially do more harm than good.

One source of unintended mutations in CRISPR is the mechanisms by which DNA repairs itself. In fact, after Cas9 cleaves the DNA in a cell, the cell’s own repair mechanisms can introduce mutations that silence critical genes.

When Cas9 produces a double-stranded break in the DNA, it is ligated back together via one of two endogenous repair mechanisms: homology-directed repair (HDR) or non-homologous end joining (NHEJ). HDR uses a repair template that is homologous to the damaged DNA to rejoin the two DNA strands, and this method typically does not produce unintended mutations.

But HDR is less common than NHEJ, and NHEJ is more efficient than HDR. Yet, NHEJ produces errors more frequently.

Unlike HDR, NHEJ does not use a repair template. Rather, NHEJ involves ligating the two loose ends of the DNA together using one or two nucleotides. Unfortunately, these short nucleotide sequences, or indels, can cause a frameshift mutation that can introduce a stop codon that ultimately silences the gene. Furthermore, in diploid cells, these mutations can be inserted in one of three zygosities: monoallelic, where one DNA strand is edited, diallelic heterozygous, where both strands are edited with non-identical mutations, and diallelic homozygous, where both strands are edited in the same fashion. Due to the randomness of NHEJ, DNA repair is one of the primary barriers to CRISPR’s specificity.

Low efficiency

Another limitation of CRISPR is its low success rate, especially in cases when multiple edits are involved. In fact, CRISPR only enters cells and makes all its edits successfully about 5 to 36 percent of the time (as noted in the paper “Rapid and highly efficient mammalian cell engineering via Cas9 protein transfection” in the Aug. 20, 2015, issue of the Journal of Biotechnology). CRISPR efficiency depends on several factors, including the cell type, the transfection method, which gene is targeted for editing and other experimental conditions.

One reason CRISPR/Cas9 fails to enter cells is because the DNA plasmids carrying the technology are too large to cross most cell membranes. Subsequently, researchers use techniques such as viral transduction and electroporation to bypass the membranes’ narrow filter.

But CRISPR/Cas9 is even too large for some viruses. Vectors such as adeno-associated virus are too small to encapsulate the CRISPR/Cas9 system, and therefore, they form their own barrier to entry for the gene editing technology.

Electroporation, on the other hand, uses an electrical current to poke holes in cell membranes, which allows CRISPR’s passage into a cell. This method, however, can damage or even kill cells. When performing electroporation, it is critical to balance several variables, including the strength of the electric field and the nature of the buffers used, to allow efficient CRISPR transfection while maintaining the cell’s viability.

Recently, researchers created several new techniques that overcome several of CRISPR limitations, including its low transfection efficiency and its tendency to introduce unintended mutations.

DNA-free CRISPR

Many of CRISPR’s specificity issues stem from its resilience in cells. Normally, CRISPR is transfected in the form of DNA plasmids encoding both Cas9 and sgRNA, but these plasmids are resistant to degradation in cells. This causes their products to be transcribed constitutively, overpopulating the cell with gene-editing machinery. This often leads to a high frequency of off-target effects and, in some cases, toxicity.

Researchers have developed two “DNA-free” CRISPR protocols that limit the expression of the CRISPR/Cas9 system and prevent it from causing a significant number of off-target mutations. DNA-free CRISPR utilizes gene products that degrade faster than DNA plasmids, enabling only transient Cas9 expression and preventing the accrual of off-target effects.

The first of these DNA-free CRISPR methods uses a ribonucleoprotein (RNP) complex composed of an sgRNA that is transcribed and complexed with Cas9 prior to transfection. After transfection, these RNPs enable gene editing, but they degrade rather quickly, limiting the incidence of off-target effects. These RNPS are efficient, but in some cell types, the large size of the RNP prevents it from entering the cell.

The second DNA-free CRISPR method solves this issue. Instead of transfecting an RNP, a researcher can transfect transcribed sgRNA along with, instead of the Cas9 protein, a Cas9 mRNA. These mRNAs are smaller than RNPs, enabling CRISPR to edit the genome without the need for viruses, electroporation and other hazardous transfection methods, thereby increasing efficiency while maintaining the cell’s health. These mRNAs degrade faster than their DNA plasmid counterparts, however. In this system, the Cas9 mRNA might even be degraded before it can be translated in the cell.

To increase the mRNA’s lifespan in a cell, it can be polyadenylated prior to transfection. The Poly(A) tail acts as a “lightning rod” for exonucleases and slows the degradation of mRNA, enabling efficient editing. A researcher can control the lifespan of the Cas9 mRNA by the length of the attached poly(A) tail: if the tail is too long, the mRNA will overexpress, causing unintended mutations, but if it is too short, the mRNA will be degraded before it can be transcribed. Capillary electrophoresis is an efficient method for analyzing the length of these poly(A) tails.

Even with these advancements in CRISPR techniques, the technology still faces several hindrances that can affect its accuracy and effectiveness. Consequently, researchers must adopt methods to screen for positively edited sequences according to the specific criteria of their experiments to ensure their edits meet their needs.

Heteroduplex cleavage assays

The typical screening process for CRISPR edits involves several time-consuming and labor-intensive steps. After a researcher screens the original cell pools, he or she sorts individual cell lines. Then, these cell lines are screened for the presence of the desired mutation, their zygosity is tested and they are sequenced to determine the exact character of their mutations. Next-generation sequencing (NGS) is the gold standard for this type of screening protocol, yet it is costly and time-consuming. Whatever methods that can be employed to reduce the amount of DNA that needs to be sequenced will reduce costs and simplify the CRISPR workflow.

A heteroduplex cleavage assay can quickly identify the zygosity of a mutation and identify properly inserted sequences, allowing researchers to preemptively screen out cells with undesired edits prior to NGS. These assays use an endonuclease such as T7 to compare alleles and identify and cleave DNA mismatches. By analyzing the length of the resulting fragments, a researcher can estimate the zygosity of a mutation. This type of assay provides a rapid screening step prior to sequencing; identifying and isolating DNA sequences with the desired zygosity will reduce the number of clones that need to be sequenced.

The next generation of CRISPR

CRISPR’s off-target effects and low efficiency must be overcome before it is used in the clinic. Using the molecular techniques mentioned here represents a new approach to CRISPR that goes a long way toward remedying the editing technology’s largest drawbacks. It is likely that CRISPR will undergo several more generations of advancement before it truly becomes the ultimate gene-editing technology it was originally intended to be.

Steve Siembieda is the vice president of commercialization at Advanced Analytical Technologies Inc., which this year launched the FEMTO Pulse, a capillary electrophoresis-based fragment analysis tool designed for long-read sequencing. Siembieda has been with AATI since 2007.

Successful genome editing is a rare outcome that depends substantially on experimental conditions including type of cells used, transfection method, target sequence and many other factors, the company notes, and current methods for assessing genome edit efficiency, including next-generation sequencing (NGS) and high-resolution melt analysis, present drawbacks in cost, time, simplicity and sensitivity.

Bio-Rad’s ddPCR technology is reportedly well suited for the task of genome editing, empowering scientists to precisely evaluate the efficiency of their experiment in less time and at lower cost than with any other method. By partitioning samples into thousands of droplets, ddPCR technology increases the signal-to-noise ratio, which allows users to quantify extremely rare edits—even frequencies of 0.5 percent and from as little as 5 ng of genomic DNA—and still obtain the results within one day.

Bio-Rad’s offering follows what it calls “a wave of research in ddPCR methods to measure genome-editing efficiency,” noting that in 2015, Science published a ddPCR-based method used at Duke University to detect edits designed to treat Duchenne muscular dystrophy in mice. The following year, research papers in Nature Protocols and Scientific Reports further detailed more ddPCR strategies for assessing genome editing outcomes. In a study released this year, researchers at the Broad Institute of MIT and Harvard used ddPCR to verify the sensitivity of an innovative CRISPR-based nucleic acid detection platform.

“Genome editing holds great promise not only in basic and applied science, but particularly in the area of gene therapy,” said Boris Fehse, a professor of cell and gene therapy in Hamburg, Germany, and an author of the Nature Protocols paper. “Therapeutic applications require reliable, highly sensitive and easy-to-perform assays to monitor efficiency as well as potential side effects. In my opinion, digital PCR represents an ideal tool fulfilling these requirements.”

Should criticism of CRISPR/Cas9 curb market enthusiasm?

As market-watchers at Leerink Partners noted June 9, “Since a controversial article appeared on the reputable Nature Methods on May 30, stocks of companies utilizing CRISPR/Cas9 genome-editing technology have suffered significant losses in share price and market cap. Since then, we had the chance to review two responses challenging the reported findings of Schaefer, KA et al. as inaccurate and misleading pending further review. Admittedly, the responses emerged from Editas and Intellia, which some could attribute as a skewed opinion, but we nevertheless continue to believe that the advancements within these companies continue to support robust sets of data supporting the clinical use of CRISPR/Cas9 technology in medicine.”

Furthermore, the firm noted, it believes the stock sell-off was overdone, and although Intellia stock, for one, had somewhat regained its losses by the time they wrote the June 9 note, Leerink continued to reiterate its Outperform rating on Intellia.

According to Leerink's view on the matter, it is unclear whether the reported off-target mutations (including insertions and deletions/indels and single-nucleotide variants, or SNVs) were truly induced by CRISPR/Cas9 editing.

“Through whole-genome sequencing (WGS), Schaefer, KA et al. reported identifying 1397 SNVs and 117 indels common to the two independently generated CRISPR-edited mice, which is an alarmingly high number,” Leerink admitted, but added that there are three issues with the authors’ assessment, however.

“First, it is difficult to conclude that an observation arising from two independent mice can constitute an overarching problem with the CRISPR/Cas9 technology. To err on the side of caution, Bothmer, A et al. recommends testing several different cohorts of mice investigating various combinations of the CRISPR/Cas9 toolkit before one can be more confident of the true origins of these indels and SNVs. Second, Schaefer, KA et al. compared the genome of the edited mice to ‘one uncorrected control.’ Unfortunately, failure to compare (or comment on) the control mouse to the parents of the two mice used in this study renders the comparison inaccurate and misleading. Related to this issue, Barnes, TM et al. cites two different studies which conclude that variations exist even among inbred mouse littermates and that Cas9 induced no expected mutations. Lastly, despite the authors’ assertion, close resemblance of the observed SNVs (in both position and nucleotide change) between the two edited mice suggest that the majority of mutations were already present before the edits. According to Bothmer, A et al., ‘the odds of the exact nucleotide changes occurring in the exact same position of the exact same gene at the exact same ratios in almost every case are effectively zero.’”

Also, Leerink noted, it is difficult to know whether the investigated guide RNA (gRNA) was indeed foolproof from inducing off-target effects, pointing out that both Bothmer, A et al. and Barnes, TM et al. raised issues with the gRNA used in this study—its “high propensity for off-target effects when analyzed under gRNA specificity prediction algorithms and that the observed mutations contradict mechanisms published in the literature,” including Watson-Crick gRNA-DNA base pairing.

“We note that Schaefer, KA et al. initially tested four sgRNAs in cells before advancing with the ‘highest activity’ candidate in vivo. In contrast, Intellia has combined informatic guide selection with accurate off-target assessments to identify no off-target activity. Whether this gRNA selection process is foolproof remains to be seen, but we are inclined to assign higher credibility to Intellia’s methods than an investigator-led four gRNA-based selection process.”

Leerink did admit that the Schaefer, KA et al. publication, “while controversial, raises an important consideration inherent in all scientific endeavors—all findings are subject to scrutiny until demonstrated otherwise.”

A 2016 survey published in Nature, the firm noted, underscores the variability in experimental outcome whether the same or a different group attempts to reproduce a finding, and going back to to Intellia add that, for their part, the company “has taken the promising technology of CRISPR/Cas9 technology and ran a series of in-vitro, in-silico and in-vivo experiments to substantiate continued investigation of this genome-editing technology. Although there is no guarantee on the outcome of CRISPR/Cas9 use in the clinic, we maintain our positive view on Intellia until proven otherwise.”

In fact, in another note on Aug. 1, Leerink remarked, “Intellia has now shown effective and precise targeting with sustained and durable efficacy across mice and rats,” and, building on previous successes in rodents, also disclosed at the beginning of August an update in the application in a new animal model: non-human primates (NHPs). Also, Leerink said of Intellia’s work, “CRISPR/Cas9-treated mice have reached their nine-month time point. No premature deaths have been reported all the while retaining high efficacy—70 percent editing and 97 percent reduction in serum transtheyretin (TTR) protein. Similarly high efficacy was reproduced in rats—66 percent editing and 91 percent reduction in serum TTR protein. With the newly disclosed NHP data (albeit using GFP expression), we believe the mounting data further supports precise on-target effects of the CRISPR/Cas9 technology without any significant off-target AEs [adverse effects] ... previously raised in the scientific literature.”

As part of the collaboration, CRISPR/Cas9 gene editing will be utilized to improve upon current T cell therapies in development, ultimately addressing unmet needs in both hematologic and solid tumors. Dr. Marcela V. Maus, director of the Cellular Immunotherapy Program at MGHCC and assistant professor of medicine at Harvard Medical, will lead the scientific work at MGH.

“It is becoming increasingly clear that CRISPR/Cas9 can play a major role in enabling the next generation of T cell therapies in oncology. By combining our gene-editing capabilities with Dr. Maus’ pioneering expertise in T cell therapy, we hope to accelerate our progress toward making these therapies a reality for patients suffering from cancer,” said Dr. Jon Terrett, head of Immuno-Oncology Research and Translation for CRISPR Therapeutics.

“We have already seen the profound benefit that T cell therapies can have for certain patients with a specific set of tumor types. Now the potential with gene editing, and specifically CRISPR/Cas9, exists to create improved versions of these cells that may work for a wider variety of patients with a more diverse set of tumor types. I’m glad to see the commitment CRISPR Therapeutics is making to this area, and am excited to collaborate with them,” said Maus.

Combining CRISPR/Cas9 with cell therapies to fight cancer

CAMBRIDGE, Mass., & MILAN, Italy—June saw Intellia Therapeutics, a leading genome editing company, and San Raffaele University and Research Hospital, a leading scientific institution, enter into a three-year research collaboration, option and license agreement to engineer optimized T cell cancer therapies.

The goal of the collaboration is to discover innovative tools to target tough-to-treat cancers, while leveraging Intellia’s proprietary CRISPR/Cas9 platform to generate next-generation T cell therapies that will address unmet needs in both hematological and solid tumors. Prof. Chiara Bonini, head of San Raffaele’s Experimental Hematology Unit and deputy director of the Division of Immunology, Transplantation and Infectious Diseases, will lead the scientific work at San Raffaele.

The collaboration marks the first external partnership of Intellia’s eXtellia division. eXtellia’s long-term strategy is focused on advancing new generations of engineered cell therapies through the unique and proprietary applications of CRISPR genome editing. eXtellia was established in 2016, and has identified its initial areas of focus as immuno-oncology and autoimmunity, where genome-edited cell therapy offers a potentially powerful and differentiated therapeutic modality. The agreement also includes options and licenses to key technologies for production of engineered cell therapies developed at San Raffaele.

“Through this collaboration, eXtellia aims to apply CRISPR/Cas9 genome editing in a multifaceted way to modulate the fundamental properties of engineered immune cells and amplify their anticancer properties far beyond current applications,” said Dr. Andrew Schiermeier, senior vice president at eXtellia.